Practical Biotechnology P Ramadass, A Wilson Aruni
INDEX
×
Chapter Notes

Save Clear


Cell CultureCHAPTER ONE

 
1.1 INTRODUCTION
Since the discovery by Enders, (1949) that polioviruses could be cultured successfully in non-neural tissue, cell culture has become a very useful and convenient method for isolating viruses in vitro. Even though more modern diagnostic virological techniques such as PCR, enzyme immunoassay (EIA) and immunofluorescence (IF) have become increasingly popular recently, viral isolation in cell culture still remains the “gold standard” for many cultivable viruses. A single cell line could be used to cultivate a broad spectrum of viral agents. Viral culture also facilitates the production of high titered virus used in antibody testing, viral characterization or molecular analysis.
Most diagnostic virology laboratories use monolayer cell cultures to propagate viruses. The main advantage of using monolayer cultures is the ease with which the infected cultures can be monitored microscopically. Many viruses present themselves in cell culture by producing degenerative changes in the cells, the so-called cytopathic effect (CPE). The CPE is often characteristic of a specific virus and this allows the experienced observer to make a presumptive diagnosis based on the type of CPE present on the monolayer. A more definite viral diagnosis is carried out by further testing of the viral isolate. This can be achieved by performing a viral neutralization assay on the isolate in fresh cell culture. A useful alternative to viral neutralization is the application of immunoassay techniques such as IF staining of infected cells, EIA or nucleic acid hybridization. The application of these techniques are particularly useful for detecting specific viral replication in cultures in the absence of a CPE. Not all viruses will produce CPE and some viruses are slow to grow. Immunoassay techniques can also allow early detection of viral replication prior to the formation of a CPE and allow more rapid viral diagnosis. The availability of specific and sensitive monoclonal antibodies directed against viral antigen has greatly enhanced the use of these techniques in viral diagnosis.2
 
1.2 BASIC TECHNIQUES—THE “DO'S AND DON'TS” OF CELL CULTURE
Given below are a few of the essential “do's and don'ts” of cell culture. Some of these are mandatory, e.g. use of personal protective equipment (PPE). Many of them are common sense and apply to all laboratory areas. However, some of them are specific to tissue culture.
 
The Do's
  1. Use personal protective equipment, (laboratory coat/gown, gloves and eye protection) at all times. In addition, thermally insulated gloves, full-face visor and splash-proof apron should be worn when handling liquid nitrogen.
  2. Always use disposable caps to cover hair.
  3. Wear dedicated PPE for tissue culture facility and keep separate from PPE worn in the general laboratory environment. The use of different coloured gowns or laboratory coats makes this easier to enforce.
  4. Keep all work surfaces free of clutter.
  5. Correctly label reagents including flasks, medium and ampoules with contents and date of preparation.
  6. Only handle one cell line at a time. This common sense point will reduce the possibility of cross-contamination by mislabeling. It will also reduce the spread of bacteria and Mycoplasma by the generation of aerosols across numerous opened media bottles and flasks in the cabinet.
  7. Clean the work surfaces with a suitable disinfectant (e.g. 70% ethanol) between operations and allow a minimum of 15 minutes between handling different cell lines.
  8. Wherever possible maintain separate bottles of media for each cell line in cultivation.
  9. Examine cultures and media daily for evidence of gross bacterial or fungal contamination. This includes medium that has been purchased commercially.
  10. Quality Control all media and reagents prior to use.
  11. Keep cardboard packaging to a minimum in all cell culture areas.
  12. Ensure that incubators, cabinets, centrifuges and microscopes are cleaned and serviced at regular intervals.
  13. Test cells for Mycoplasma on a regular basis.
3
 
The Don'ts
  1. Do not continuously use antibiotics in culture medium as this will inevitably lead to the appearance of antibiotic resistant strains and may render a cell line useless for commercial purposes.
  2. Do not allow waste to accummulate particularly within the microbiological safety cabinet or in the incubators.
  3. Do not have too many people in the lab at any one time.
  4. Do not handle cells from unauthenticated sources in the main cell culture suite. They should be handled in quarantine until quality control checks are complete.
  5. Avoid keeping cell lines continually in culture without returning to frozen stock.
  6. Avoid cell culture becoming fully confluent. Always sub-culture at 70–80%.
  7. Do not allow media to go out of date. Shelf-life is only 6 weeks at +4°C once glutamine and serum are added.
  8. Avoid water baths from becoming dirty by using sanitser
  9. Do not allow essential equipment to become out of calibration. Ensure microbiological safety cabinets are tested regularly.
 
1.3 MATERIALS
  • Tissue culture flasks (Costar, Nunc, Falcon)
  • Microtiter plates, cell clusters (Costar, Nunc, Falcon)
  • U-bottomed cell culture microtiter plates (Costar)
  • Multichannel pipette
  • Glass tissue culture tubes
  • Silicon rubber bungs
  • Cell counting chamber
  • Cell cultures
The following reagents can be obtained from Gibco BRL:
  • Minimum essential medium (MEM) with Earle's salts without L-glutamine and sodium bicarbonate (10x)
  • L-glutamine 200 mM (10x)
  • MEM nonessential amino acids (100x)
  • Trypsin-EDTA (1x)
  • Penicillin-streptomycin solution (10,000 Units)
  • Gentamycin sulfate (10 mg/ml)
  • Fungizone (250 mg/ml)
  • Sodium bicarbonate
    4
  • Fetal bovine serum
  • Nystatin (100x), 10,000 Units/ml
  • Hanks balanced salt solution (10x)
  • PBS tablets.
 
1.4 TRANSPORT MEDIUM
Virus viability is crucial for successful isolation in cell culture. Enveloped viruses are particularly labile. Viruses, such as respiratory syncytial virus, influenza virus and the herpes virus, may lose infectivity if they are not adequately protected. Rapid transportation to the laboratory under the proper conditions can greatly enhance effective isolation. Viruses should be transported to the laboratory in the appropriate transport medium (viral transport medium) which can be bought commercially or made up in-house.
 
1.5 VIRAL TRANSPORT MEDIUM (500 ml)
Hanks balanced salt solution (10x)
50 ml
Distilled water, Millipore
415 ml
Fetal bovine serum
10 ml
Penicillin/streptomycin
10 ml
Gentamycin sulfate
2.5 ml
Fungizone
2.5 ml
Few drops of sodium hydroxide, 1 M
pH 7.2-7.4
HEPES buffer
10 ml
Made up as described and aseptically dispensed into 2 ml aliquots.
 
1.6 2X EMEM MAINTENANCE MEDIUM (100 ml)
Twenty ml 10x MEM; 68 ml sterile water; 2 ml penicillin/streptomycin; 0.4 ml nystatin (50 U/ml). Just before use, 8 ml of sodium bicarbonate (7.5%, w/v) and 2 ml fetal calf serum (pH 7.4) were added.
 
1.7 GROWTH MEDIUM (500 ml stock)
Stock solution was prepared as described and stored at 4°C until required. Sodium bicarbonate and fetal bovine serum are not added to the stock solution until it is about to be used.
MEM (10x)
50 ml
Sterile water
433 ml
Non-essential amino acids (NEAA) (100x)
5 ml
5
L-glutamine
5 ml
Penicillin-streptomycin
5 ml
Fungizone
2 ml
Sodium bicarbonate 7.5%
Use at 2–3 ml/100 ml stock
Fetal bovine serum
Use at 10%
Maintenance medium (MM) (500 ml stock)
MEM (10x)
50 ml
Sterile water
438 ml
NEAA (100x)
5 ml
Penicillin-streptomycin
5 ml
Fungizone
2 ml
Sodium bicarbonate 7.5%
Use at 4 ml/100 ml stock
Fetal bovine serum
Use at 1%
Table 1.1   Different types of culture media and their uses
Media type
Examples
Uses
Balanced salt solutions (BSS)
PBS, Hanks BSS, Earle's salts, Dulbecco's PBS, HBSS and EBSS
Form the basis of many complex media
Basal media
MEM
Primary and diploid cultures
DMEM
Modification of MEM containing increased level of amino acids and vitamins. Supports a wide range of cell types including hybridomas.
GMEM
Glassgow's modified MEM was defined for BHK-21 cells
Complex media
RPMI 1640 (Roswell Park Memorial Institute)
Originally derived from human leukemic cells. It supports a wide range of mammalian cells including hybridomas
Iscoves DMEM
Further enriched modification of DMEM which supports high density growth
TC 100, Grace's insect medium, Schenider's insect medium, Mitsuhasi Moromosh medium
Designed for culturing baculovirus in lepidopteron cell lines (such as Spodoptera frugiperda
Serum free media
CHO (Chinese hamster ovary cells)
For use in serum-free applications
Ham F-10 DMEM/F-12
These media are usually HEPES buffered
Insect cells
Sf-900 II SFM, SF insect medium-2
Specifically designed for use with Sf 9 insect cells
6
 
1.8 TISSUE CULTURE MEDIA
We use two different kinds of media. Most cells are grown in DMEM. A few lymphoid cell lines are grown in RPMI. Cells grown in DMEM must be grown in a 10% CO2 atmosphere. In contrast, cells grown in RPMI must be grown in a 5% CO2 atmosphere. If cells are grown in RPMI in 10% CO2, the medium will be too acidic (yellow).
 
1.9 DMEM
This is an abbreviation for Dulbecco/Vogt modified Eagle's (Harry Eagle) minimal essential medium. We use commercially-made DMEM as the medium for most cells. DMEM differs from the original MEM in that it contains approximately 4 times as much of the vitamins and amino acids present in the original formula, and some (2 to 4 fold) more glucose. It also contains iron and a few other oddments. Most kinds of cells—human, monkey, hamster, rat, mouse, chicken—grow well in this medium. Different types of culture media and their uses are given in Table 1.1.
 
1.10 RPMI
This is an abbreviation for Roswell Park Memorial Institute medium. Human lymphoid cells are traditionally grown in RPMI medium. This medium contains a great deal of phosphate and is formulated for 5% CO2. Medium is always supplemented with serum. Most cell lines are grown in DMEM supplemented with either 10% calf serum or 10% fetal calf serum. Transformed cells usually grow well in calf serum. Normal (non-transformed) cells sometimes require fetal calf serum, which is richer. Some clones of 3T3 cells grow better in fetal calf serum. Some very fastidious cells are grown in 20% fetal calf serum.
Due to the cost and scarcity of fetal calf serum, some cells are grown in horse serum instead. The medium of some lymphoid cells is supplemented with 5 × 10−5 M 2-mercaptoethanol. It does not matter whether this has undergone oxidation.
Some cells benefit from the inclusion of 2 mM glutamine, 2, 5 or 10% tryptose phosphate broth, 2 mM pyruvate, or non-essential amino acids in their media.
 
1.11 BASIC CONSTITUENTS OF MEDIA
  • Inorganic salts
  • Carbohydrates
    7
  • Amino acids
  • Vitamins
  • Fatty acids and lipids
  • Proteins and peptides
  • Serum
Each type of constituent performs a specific function as outlined below:
 
Inorganic Salts
The inclusion of inorganic salts in media performs several functions. Primarily they help to retain the osmotic balance of the cells and help to regulate membrane potential by provision of sodium, potassium and calcium ions. All of these are required in the cell matrix for cell attachment and as enzyme cofactors.
 
Buffering Systems
Most cells require pH conditions in the range 7.2 to 7.4 and close control of pH is essential for optimum culture conditions. There are major variations to this optimum. Fibroblasts prefer a higher pH (7.4-7.7) whereas, continuous transformed cell lines require more acid conditions pH (7.0-7.4). Regulation of pH is particularly important immediately following cell seeding when a new culture is established and is usually achieved by one of the two buffering systems; (i) a “natural” buffering system where gaseous CO2 balances with the CO3/HCO3 content of the culture medium and (ii) chemical buffering using a zwitterion called HEPES.
Cultures using natural bicarbonate/CO2 buffering systems need to be maintained in an atmosphere of 5 to 10% CO2 in air usually supplied in a CO2 incubator. Bicarbonate/CO2 is low cost, non-toxic and also provides other chemical benefits to the cells. HEPES has superior buffering capacity in the pH range 7.2 to 7.4 but is relatively expensive and can be toxic to some cell types at higher concentrations. HEPES buffered cultures do not require a controlled gaseous atmosphere. Most commercial culture media include phenol red as a pH indicator so that the pH status of the medium is constantly indicated by the colour. Usually the culture medium should be changed/replenished if the colour turns yellow (acid) or purple (alkali).
 
Carbohydrates
The main source of energy is derived from carbohydrates generally in the form of sugars. The major sugars used are glucose and galactose, however some media contain maltose or fructose. The concentration of sugar varies 8from basal media containing 1 g/L to 4.5 g/L in some more complex media. Media containing the higher concentration of sugars are able to support the growth of a wider range of cell types.
 
Vitamins
Serum is an important source of vitamins in cell culture. However, many media are also enriched with vitamins making them consistently more suitable for a wider range of cell lines. Vitamins are precursors for numerous co-factors. Many vitamins, especially B group vitamins are necessary for cell growth and proliferation and for some cell lines the presence of B12 is essential. Some media also have increased levels of vitamins A and E. The vitamins commonly used in media include riboflavin, thiamine and biotin.
 
Proteins and Peptides
These are particularly important in serum-free media. The most common proteins and peptides include albumin, transferrin, fibronectin and fetuin and are used to replace those normally present through the addition of serum to the medium.
 
Fatty Acids and Lipids
Like proteins and peptides these are important in serum-free media since they are normally present in serum, e.g. cholesterol and steroids are essential for specialized cells.
 
Trace Elements
These include trace elements such as zinc, copper, selenium and tricarboxylic acid intermediates. Selenium is a detoxifier and helps remove oxygen free radicals. Whilst all media may be made from the basic ingredients, this is time consuming and may predispose to contamination. For convenience most media are available as ready mixed powders or as 10x and 1x liquid media. If powder or 10x media are purchased it is essential that the water used to reconstitute the powder or dilute the concentrated liquid is free from mineral, organic and microbial contaminants. It must also be pyrogen-free. In most cases water prepared by reverse osmosis and resin cartridge purification with a final resistance of 16 to 18 Mx is suitable. Once prepared, the media should be filter sterilized before use. Obviously, purchasing 1x liquid media eliminates the need for this.9
 
Serum
Serum is a complex mix of albumins, growth factors and growth inhibitors and is probably one of the most important components of cell culture medium. The most commonly used serum is fetal bovine serum. Other types of serum are available including newborn calf serum and horse serum. The quality, type and concentration of serum can all affect the growth of cells and it is therefore important to screen batches of serum for their ability to support the growth of cells. In addition there are other tests that may be used to aid the selection of a batch of serum including cloning efficiency, plating efficiency and the preservation of cell characteristics.
Serum is also able to increase the buffering capacity of cultures that can be important for slow growing cells or where the seeding density is low (e.g. cell cloning experiments). It also helps to protect against mechanical damage which may occur in stirred cultures or whilst using a cell scraper. A further advantage of serum is the wide range cell types with which it can be used despite the varying requirements of different cultures in terms of growth factors. In addition, serum is able to bind and neutralize toxins. However, serum is subject to batch-batch variation that makes standardization of production protocols difficult. There is also a risk of contamination associated with the use of serum. These risks can be minimized by obtaining serum from a reputable source since suppliers of large quantities of serum perform a battery of quality control tests and supply a certificate of analysis with the serum. In particular, serum is screened for the presence of bovine viral diarrhea virus (BVDV) and Mycoplasma. Heat inactivation of serum (incubation at 56°C for 30 minutes) can help to reduce the risk of contamination since some viruses are inactivated by this process. However, the routine use of heat inactivated serum is not an absolute requirement for cell culture. The use of serum also has a cost implication not only in terms of medium formulation but also in downstream processing. A 10% Fetal bovine serum (FBS) supplement contributes 4.8 mg of protein per milliliter of culture fluid, which complicates downstream processing procedures.
 
1.12 GUIDELINES FOR SERUM USE
Fetal bovine serum (FBS) has been used to prepare a number of biologicals and has an excellent record of safety. The recognition of Bovine spongiform encephalopathy (BSE) in 1986 and its subsequent spread into continental Europe along side the announcement of the probable link between BSE and a new variant of Creutzfeldt-Jacob disease in humans, stimulated an increased concern about safe sourcing of all bovine materials. In 1993, the Food and Drug Administration (FDA) recommended against the use of bovine derived materials from cattle which have resided in, or originated from countries where BSE has been diagnosed. The current European Union (EU) guidelines on 10viral safety focus on sourcing, testing and paying particular attention to the potential risk of cross-contamination during slaughtering or collection of the starting tissue.
The use of FBS in production processes of medicinal products is acceptable, provided good documentation on sourcing, age of the animals and testing for the absence of adventitious agents is submitted. All responsible suppliers of FBS for bio-pharmaceutical applications will provide such documentation.
Recent regulatory requirements in Europe stress the importance of justifying the use of material of bovine, caprine or ovine origin in the production of pharmaceutical products. Thus, although FBS has been used for many years in the production process of many medicinal products such as viral vaccines and recombinant DNA products, at present there is a justified trend to remove all material of animal origin from manufacturing processes.
 
1.13 PROTOCOL-1: ASEPTIC TECHNIQUE AND GOOD CELL CULTURE PRACTICE
 
Aim
To ensure all cell culture procedures are performed to a standard that will prevent contamination from bacteria, fungi and Mycoplasma and cross- contamination with other cell lines.
 
Reagents and Equipment
  • Sanitisers
  • 1% Formaldehyde-based disinfectant
  • 70% Ethanol in water
  • Personal protective equipment (sterile gloves, laboratory coat, safety visor)
  • Microbiological safety cabinet at appropriate containment level.
 
Procedure
  1. Sanitize the cabinet using 70% ethanol before commencing work.
  2. Sanitize gloves by washing them in 70% ethanol and allowing to air dry for 30 seconds before commencing work.
  3. Put all materials and equipment into the cabinet prior to starting work after sanitizing the exterior surfaces with 70% ethanol.
  4. Whilst working do not contaminate gloves by touching anything outside the cabinet (especially face and hair). If gloves become contaminated re-sanitize with 70% ethanol as above before proceeding.
  5. Discard gloves after handling contaminated cultures and at the end of all cell culture procedures.
  6. Equipment in the cabinet or that which will be taken into the cabinet during cell culture procedures (media bottles, pipette tip boxes, pipette 11aids) should be wiped with tissue soaked with 70% ethanol prior to use.
  7. Movement within and immediately outside the cabinet must not be rapid. Slow movement will allow the air within the cabinet to circulate properly.
  8. Speech, sneezing and coughing must be directed away from the cabinet so as not to disrupt the airflow.
  9. After completing work, disinfect all equipment and material before removing from the cabinet. Spray the work surfaces inside the cabinet with 70% ethanol and wipe dry with tissue. Dispose of tissue by autoclaving.
  10. Cell culture discard in chloros (10,000 ppm) must be kept in the cabinet for a minimum of two hours (preferably overnight) prior to discarding down the sink with copious amounts of water.
  11. Periodically clean the cabinet surfaces with a disinfectant or fumigate the cabinet according to the manufacturers instructions. However, you must ensure that it is safe to fumigate your own laboratory environment due to the generation of gaseous formaldehyde.
 
1.14 PROTOCOL-2: RESUSCITATION OF FROZEN CELL LINES
 
Aim
Many cultures obtained from a culture collection, such as ATCC, ECACC, will arrive frozen and in order to use them the cells must be thawed and put into culture. It is vital to thaw cells correctly in order to maintain the viability of the culture and enable the culture to recover more quickly. Some cryoprotectants, such as DMSO, are toxic above 4°C. Therefore it is essential that cultures are thawed quickly and diluted in culture medium to minimize the toxic effects.
 
Reagents and Equipment
  • Media—prewarmed to the appropriate temperature (refer to the Cell Line Data Sheet sent along with the cell line for the correct medium and size of flask to resuscitation into)
  • 70% Ethanol in water
  • DMSO
  • Personal protective equipment (sterile gloves, laboratory coat, safety visor)
  • Waterbath set to appropriate temperature
  • Microbiological safety cabinet at appropriate containment level
  • CO2 incubator
  • Pre-labeled flasks
  • Pipettes
  • Ampoule rack
  • Tissue paper
12
 
Procedure
  1. Read technical data sheet to establish specific requirements for the cell line.
  2. Prepare the flasks as appropriate (information on technical data sheet). Label with cell line name, passage number and date.
  3. Collect ampoule of cells from liquid nitrogen storage wearing appropriate protective equipment and transfer to laboratory in a sealed container.
  4. Still wearing protective clothing, remove ampoule from container and place in a waterbath at an appropriate temperature for the cell line, e.g. 37°C for mammalian cells. Submerge only the lower half of the ampoule. Allow to thaw until a small amount of ice remains in the vial-usually 1–2 minutes. Transfer to class II safety cabinet.
  5. Wipe the outside of the ampoule with a tissue moistened (not excessively) with 70% alcohol and hold tissue over ampoule to loosen lid.
  6. Slowly, dropwise, pipette cells into prewarmed growth medium to dilute out the DMSO (flasks prepared in step 2).
  7. Incubate at the appropriate temperature for species and appropriate concentration of CO2 in atmosphere.
  8. Examine cells microscopically (phase contrast) after 24 hours and sub-culture as necessary.
 
Key Points
  1. Most textbooks recommend washing the thawed cells in media to remove the cryoprotectant. This is only necessary if the cryoprotectant is known to have an adverse effect on the cells. In such cases the cells should be washed in media before being added to their final culture flasks.
  2. Do not use an incubator to thaw cell cultures since the rate of thawing achieved is too slow resulting in a loss of viability.
  3. If a CO2 incubator is not available, gas the flasks for 1–2 minutes with 5% CO2 in 95% air filtered through a 0.25 μ filter.
  4. For some cultures it is necessary to subculture before confluence is reached in order to maintain their characteristics, e.g. the contact inhibition of 3T3 cell line is lost if they are allowed to reach confluence repeatedly.
 
1.15 PROTOCOL-3: SUBCULTURE OF ADHERENT CELL LINES
 
Aim
Adherent cell lines will grow in vitro until they have covered the surface area available or the medium is depleted of nutrients. At this point the cell lines should be subcultured in order to prevent the culture dying. To subculture the cells they need to be brought into suspension. The degree of adhesion varies from cell line to cell line but in the majority of cases proteases, e.g. trypsin, are used to release the cells from the flask. However, this may not be appropriate for some lines where exposure to proteases is harmful or where 13the enzymes used remove membrane markers/receptors of interest. In these cases cells should be brought into suspension into a small volume of medium mechanically with the aid of cell scrapers.
 
Reagents and Equipment
  • Media—prewarmed to 37°C
  • 70% Ethanol in water
  • PBS without Ca2+/Mg2+
  • 0.25% Trypsin/EDTA in HBSS, without Ca2+/Mg2+
  • Trypsin
  • Soybean trypsin inhibitor
  • Personal protective equipment (sterile gloves, laboratory coat, safety visor)
  • Waterbath set to appropriate temperature
  • Microbiological safety cabinet at appropriate containment level
  • CO2 incubator
  • Pre-labeled flasks
  • Pipettes
  • Ampoule rack
  • Tissue paper
 
Procedure
  1. View cultures using an inverted microscope to assess the degree of confluency and confirm the absence of bacterial and fungal contaminants.
  2. Remove spent medium.
  3. Wash the cell monolayer with PBS without Ca2+/Mg2+ using a volume equivalent to half the volume of culture medium. Repeat this wash step if the cells are known to adhere strongly.
  4. Pipette trypsin/EDTA onto the washed cell monolayer using 1 ml per 25 cm2 of surface area. Rotate flask to cover the monolayer with trypsin. Decant the excess trypsin.
  5. Return flask to the incubator and leave for 2 to 10 minutes.
  6. Examine the cells using an inverted microscope to ensure that all the cells are detached and floating. The side of the flasks may be gently tapped to release any remaining attached cells.
  7. Resuspend the cells in a small volume of fresh serum-containing medium to inactivate the trypsin. Remove 100–200 μl and perform a cell count (see Protocol 7- Cell Quantification).
  8. Transfer the required number of cells to a new labeled flask containing pre-warmed medium.
  9. Incubate as appropriate for the cell line.
  10. Repeat this process as demanded by the growth characteristics of the cell line.
14
 
Key Points
  1. Some cultures whilst growing as attached lines adhere only lightly to the flask, thus it is important to ensure that the culture medium is retained and the flasks are handled with care to prevent the cells detaching prematurely.
  2. Although most cells will detach in the presence of trypsin alone, the EDTA is added to enhance the activity of the enzyme.
  3. Trypsin is inactivated in the presence of serum. Therefore, it is essential to remove all traces of serum from the culture medium by washing the monolayer of cells with PBS without Ca2+/Mg2+.
  4. Cells should only be exposed to trypsin/EDTA long enough to detach cells. Prolonged exposure could damage surface receptors.
  5. Trypsin should be neutralized with serum prior to seeding cells into new flasks otherwise cells will not attach.
  6. Trypsin may also be neutralized by the addition of soybean trypsin inhibitor, where an equal volume of inhibitor at a concentration of 1 mg/ml is added to the trypsinized cells. The cells are then centrifuged, resuspended in fresh culture medium and counted as above. This is especially necessary for serum-free cell culture.
  7. If a CO2 incubator is not available, gas the flasks for 1–2 min with 5% CO2 in 95% air filtered through a 0.25 μ filter.
 
1.16 PROTOCOL-4: ESTABLISHMENT OF A PRIMARY CULTURE
 
Reagents and Equipment
  • Chick embryo (approximately 8 days old)
  • 70% (v/v) Ethanol for swabbing
  • Sterile scissors, forceps and probes
  • Sterile Petri plates
  • Phosphate buffered saline (PBS)
  • Trypsin, cold sterilized in a 125 ml sterile Erlenmeyer flask containing a magnetic stirring bar
  • Minimum essential medium
  • Fetal calf serum
  • Clinical centrifuge with sterile capped centrifuge tubes
  • Culture flasks
  • Inverted phase contrast microscope (optional).
 
Procedure
  1. Candle an 8-day-old egg to ensure that it is alive. This is easily accomplished by holding the egg in front of a bright light source; the embryo can be seen as a shadow. Circle the embryo with a pencil.
    15
  2. Place the egg in a beaker with the blunt end up, and wash the top with a mild detergent, followed by swabbing with ethanol.
  3. Carefully puncture the top of the egg with the point of a pair of sterile scissors and cut away a circle of shell, thus exposing the underlying membrane (the chorioallantois).
  4. With a second pair of sterile scissors, carefully cut away and remove the chorioallantoic membrane, exposing the embryo.
  5. Identify and carefully remove the embryo by the neck, using a sterile metal hook or a bent glass rod, and place the embryo in a 100 mm Petri dish containing phosphate buffered saline (PBS). Wash several times with PBS by transferring the embryo to fresh Petri plates. After removal of all yolk and/or blood, transfer the embryo to a clean dish with PBS.
  6. Using two sterile forceps, remove the head, limbs, and viscera. Be sure to remove the entire limb by pulling at the proximal end. Move the remaining tissues of the embryo to yet another dish and wash with PBS.
  7. Mince the embryo finely with scissors and transfer the minced tissue to a flask containing PBS. Allow the tissue pieces to settle.
  8. Remove the PBS with a sterile pipette and add 25 ml of trypsin, a proteolytic enzyme. Stir the solution gently at 37° C for 15–20 minutes.
  9. Allow the larger, undigested tissue pieces to settle and decant the supernatant into an equal volume of minimal essential medium (MEM) + 10% fetal calf serum (FCS). FCS contains protease inhibitors which will inactivate the trypsin.
  10. Centrifuge the cells in MEM at 1,000 rpm for 10 minutes in a standard clinical centrifuge. Remove the supernatant and resuspend the pellet in 25 ml of fresh MEM + 10% FCS.
  11. Remove 0.1 ml of the culture and determine the cell concentration and viability as given in the previous section.
  12. Seed two 25 cm2 plastic culture flasks containing 25 ml of MEM + 10% FCS to a final concentration of 105 cells/ml.
  13. Label and place cultures in the tissue culture incubator at 37° C and examine daily for cell density and morphology.
  14. Note any changes in the color of the media. Tissue culture media has a pH indicator (Phenol Red) added in order to check on the growth of cells. The media initially is a cherry red (with slight blue haze) and turns orange and then yellow as the cells grow, thereby reducing the media. Should this color change occur within 24 hours, the culture is most likely contaminated and should be disposed of.
  15. Examine the cultures using an inverted phase contrast microscope. This will allow observation of the cells without opening or disturbing the growth.
  16. Make cell density determinations at 10X magnification using a square ocular grid.
  17. Plot the cell density on a log scale vs time of culture.
16
 
1.17 PROTOCOL- 5: SUBCULTURE OF SEMI-ADHERENT CELL LINES
 
Aim
Some cultures grow as a mixed population (e.g. B95-8-marmoset) where a proportion of cells do not attach to the tissue culture flask and remain in suspension. Therefore to maintain this heterogeneity both the attached cells and the cells in suspension must be subcultured (Fig. 1.1).
 
Reagents and Equipment
  • Media—prewarmed to 37°C
  • 70% Ethanol in water
  • PBS without Ca2+/Mg2+
  • 0.25% Trypsin/EDTA in HBSS, without Ca2+/Mg2+
  • Trypsin
  • Soybean trypsin inhibitor
  • Personal protective equipment (sterile gloves, laboratory coat, safety visor)
  • Waterbath set to 37°C
  • Microbiological safety cabinet at the appropriate containment level
  • Centrifuge
  • Inverted phase contrast microscope
  • CO2 incubator
  • Hemocytometer (Bright-line, Improved Neubauer)
  • Pre-labeled flasks
  • Tissue paper.
zoom view
FIGURE 1.1: Adherent cell line - Vero (African Green Monkey)
17
 
Procedure
  1. View cultures using an inverted phase contrast microscope to assess the degree of confluency and confirm the absence of bacterial and fungal contaminants. Give the flask a gentle knock first, this may dislodge the cells from the flask and remove the need for a trypsinization step with the subsequent loss of some cells due to the washings.
  2. Decant spent medium into a sterile centrifuge tube and retain.
  3. Wash any remaining attached cells with PBS without Ca2+/Mg2+ using 1–2 ml for each 25 cm2 of surface area. Retain the washings.
  4. Pipette trypsin/EDTA onto the washed cell monolayer using 1 ml per 25 cm2 of surface area. Rotate flask to cover the monolayer with trypsin. Decant the excess trypsin.
  5. Return flask to an incubator and leave for 2–10 minutes.
  6. Examine the cells using an inverted microscope to ensure that all the cells are detached and floating. The side of the flasks may be gently tapped to release any remaining attached cells.
  7. Transfer the cells into the centrifuge tube containing the retained spent medium and cells.
  8. Centrifuge the remaining cell suspension at 150 g for 5 minutes. Also centrifuge the washings from step No 3 above, if they contain significant numbers of cells.
  9. Decant the supernatants and resuspend the cell pellet in a small volume (10–20 ml) of fresh culture medium. Pool the cell suspensions. Count the cells.
  10. Pipette the required number of cells to a new labeled flask and dilute to the required volume using fresh medium.
  11. Repeat this process every 2–3 days as necessary.
 
Key Points
  1. Although most cells will detach in the presence of trypsin alone the inclusion of EDTA is used to enhance the activity of the enzyme.
  2. Trypsin is inactivated in the presence of serum. Therefore, it is essential to remove all traces of serum from the culture medium by washing the monolayer of cells with PBS without Ca2+/Mg2+. Repeated warming to 37°C also inactivates trypsin.
  3. Cells should only be exposed to trypsin/EDTA long enough to detach cells. Prolonged exposure could damage surface receptors. In general a shorter time of exposure to trypsin is required for semi-adherent cell lines.
  4. Trypsin should be neutralized with serum prior to seeding cells into new flasks otherwise cells will not attach.
  5. Trypsin may also be neutralized by the addition of soybean trypsin inhibitor, where an equal volume of inhibitor at a concentration of 1 mg/ml is added 18to the trypsinized cells. The cells are then centrifuged, resuspended in fresh culture medium and counted as above.
  6. If a CO2 incubator is not available, gas the flasks for 1–2 minutes with 5% CO2 in 95% air filtered through a 0.25 μ filter.
 
1.18 PROTOCOL-6: SUBCULTURE OF SUSPENSION CELL LINES
 
Aim
In general terms cultures derived from blood (e.g. lymphocytes) grow in suspension. Cells may grow as single cells or in clumps (e.g. EBV transformed lymphoblastoid cell lines). For these types of lines subculture by dilution is relatively easy. But for lines that grow in clumps, it may be necessary to bring the cells into a single cell suspension by centrifugation and resuspension by pipetting in a smaller volume before counting (Fig. 1.2).
 
Reagents and Equipment
  • Media—prewarmed to 37°C
  • 70% Ethanol in water
  • Personal protective equipment (sterile gloves, laboratory coat, safety visor)
  • Waterbath set to 37°C
  • Microbiological safety cabinet at appropriate containment level
  • Centrifuge
  • CO2 incubator
  • Inverted phase contrast microscope
  • Hemocytometer (Bright-line, Improved Neubauer)
  • Pre-labeled flasks.
zoom view
FIGURE 1.2: Suspension cell line – B lymphoblastoma cell line
19
 
Procedure
  1. View cultures using an inverted phase contrast microscope. Cells growing in exponential growth phase should be bright, round and refractile. Hybridomas may be very sticky and require a gentle knock to the flask to detach the cells. EBV transformed cells can grow in very large clumps that are very difficult to count and the center of the large clumps may be non-viable.
  2. Do not centrifuge the subculture unless the pH of the medium is acidic (phenol red = yellow) which indicates the cells have overgrown and may not recover. If this is so, centrifuge at 150 g for 5 minutes, re-seed at a slightly higher cell density and add 10 to 20% of conditioned medium (supernatant) to the fresh media.
  3. Take a small sample of the cells from the cell suspension (100–200 μl) (Protocol 7–Cell Quantification). Calculate cells/ml and re-seed the desired number of cells into freshly prepared flasks without centrifugation just by diluting the cells. The data sheet will give the recommended seeding densities.
  4. Repeat this every 2–3 days.
 
Key Point
If the cell line is a hybridoma or other cell line that produces a substance (e.g. recombinant protein or growth factor) of interest retain the spent media for analysis.
 
1.19 PROTOCOL-7: CELL QUANTIFICATION
 
Aim
For the majority of manipulations using cell cultures, such as transfections, cell fusion techniques, cryopreservation and subculture routines it is necessary to quantify the number of cells prior to use. Using a consistent number of cells will maintain optimum growth and also help to standardize procedures using cell cultures. This in turn gives results with better reproducibility.
 
Reagents and Equipment
  • Media—prewarmed to appropriate temperature
  • 70% Ethanol in water
  • 0.4% Trypan blue solution
  • Trypsin/EDTA
  • Personal protective equipment (sterile gloves, laboratory coat, safety visor)
  • Waterbath set to appropriate temperature
  • Microbiological safety cabinet at appropriate containment level
    20
  • Centrifuge
  • CO2 incubator
  • Hemocytometer (Bright-line, Improved Neubauer)
  • Inverted phase contrast microscope
  • Pre-labeled flasks.
 
Procedure
  1. Bring adherent and semi-adherent cells into suspension using trypsin/EDTA as above (Protocol 3 and 4) and resuspend in a volume of fresh medium at least equivalent to the volume of trypsin. For cells that grow in clumps centrifuge and resuspend in a small volume and gently pipette to break up clumps.
  2. Under sterile conditions remove 100–200 μl of cell suspension.
  3. Add an equal volume of trypan blue (dilution factor =2) and mix by gentle pipetting.
  4. Clean the hemocytometer.
  5. Moisten the cover slip with water or exhaled breath. Slide the cover slip over the chamber back and forth using slight pressure until Newton's refraction rings appear (Newton's refraction rings are seen as rainbow-like rings under the cover slip).
  6. Fill both sides of the chamber (approx. 5–10 μl) with cell suspension and view under a light microscope using 20X magnification.
  7. Count the number of viable (seen as bright cells) and non-viable cells (stained blue). Ideally >100 cells should be counted in order to increase the accuracy of the cell count. Note the number of squares counted to obtain your count of >100.
  8. Calculate the concentration of viable and non-viable cells and the percentage of viable cells using the equations below.
 
Where:
  • A is the mean number of viable cells counted, i.e. total viable cells counted divided by number of squares
  • B is the mean number of non-viable cell counted, i.e. total non-viable cells counted divided by number of squares
  • C is the dilution factor and
  • D is the correction factor (this is provided by the hemocytometer manufacturer).
Concentration of viable cells (cells/ml) = A × C × D
Concentration of non-viable cells (cells/ml) = B × C × D
Total number of viable cells = concentration of viable cells × volume
Total number of cells = number of viable cells + number of dead cells
zoom view
21
 
Key Points
  1. Trypan blue is toxic and is a potential carcinogen. Protective clothing, gloves and face/eye protection should be worn. Do not breathe the vapor.
  2. The central area of the counting chamber is 1 mm2. This area is subdivided into 25 smaller squares (1/25 mm2). Each of these is surrounded by triple lines and is then further divided into 16 (1/400 mm2). The depth of the chamber is 0.1 mm.
  3. The correction factor of 104 converts 0.1 mm3 to 1 ml (0.1 mm3 = 1 mm2 × 0.1 mm).
  4. There are several sources of inaccuracy:
    • The presence of air bubbles and debris in the chamber.
    • Overfilling the chamber such that sample runs into the channels or the other chamber.
    • Incomplete filling of the chamber.
    • Cells not evenly distributed throughout the chamber.
    • Too few cells to count. This can be overcome by centrifuging the cells, resuspending in a smaller volume and recounting.
    • Too many cells to count. This can be overcome by using a higher dilution factor in trypan blue, e.g. 1:10.
 
1.20 PROTOCOL-8: CRYOPRESERVATION OF CELL LINES
 
Aim
The protocol below describes the use of passive methods involving an −80°C deep freezer for the cryopreservation of cell cultures. A programmable freezer is the most reliable and reproducible way to freeze cells but as the cost of such equipment is beyond the reach of majority of research laboratories the methods below are described in detail. If large numbers of cell cultures are regularly being frozen then a programmable freezer is recommended.
 
Reagents and Equipment
  • Freeze medium (commonly 70% basal medium, 20% FCS, 10% DMSO or glycerol)
  • 70% Ethanol in water
  • PBS without Ca2+/Mg2+
  • 0.25% Trypsin/EDTA in HBSS, without Ca2+/Mg2+
  • DMSO
  • Trypsin/EDTA
  • BHK-21 Cell line
  • Personal protective equipment (sterile gloves, laboratory coat)
  • Full-face protective mask/visor
  • Waterbath set to 37°C
    22
  • Microbiological safety cabinet at appropriate containment level
  • Centrifuge
  • Hemocytometer (Sigma Bright-line, Improved Neubauer)
  • Pre-labeled ampoules/cryotubes
  • Cell freezing device (e.g. Nalgene).
 
Procedure
  1. View cultures using an inverted microscope to assess the degree of cell density and confirm the absence of bacterial and fungal contaminants.
  2. Bring adherent and semi-adherent cells into suspension using trypsin/EDTA as above (Protocol 3 and 5 – Subculture of adherent/attached and semi-adherent cell lines) and re-suspend in a volume of fresh medium at least equivalent to the volume of trypsin. Suspension cell lines can be used directly.
  3. Remove a small aliquot of cells (100–200 μl) and perform a cell count. Ideally the cell viability should be in excess of 90% in order to achieve a good recovery after freezing.
  4. Centrifuge the remaining culture at 150 g for 5 minutes.
  5. Re-suspend cells at a concentration of 2–4×106 cells per ml in freeze medium.
  6. Pipette 1 ml aliquots of cells into cyroprotective ampoules that have been labeled with the cell line name, passage number, cell concentration and date.
  7. Place ampoules inside a passive freezer (Nalgene). Fill freezer with isopropyl alcohol and place at −80°C overnight.
  8. Frozen ampoules should be transferred to the vapor phase of a liquid nitrogen storage vessel and the locations recorded.
 
Key Points
  1. The most commonly used cryoprotectant is dimethyl sulphoxide. However, this is not appropriate for all cell lines, e.g. BHK-21, where DMSO is used to induce differentiation. In such cases an alternative such as glycerol should be used.
  2. The freeze medium recommended above has been shown to be a good universal medium for most cell types. Another commonly used freeze medium formulation is 70% basal medium, 20% FCS, 10% DMSO but this may not be suitable for all cell types. Check whether it works for your cells before using on a regular basis.
  3. It is essential that cultures are healthy and in the log phase of growth. This can be achieved by using pre-confluent cultures (cultures that are below their maximum cell density) and by changing the culture medium 24 hours before freezing.
  4. The rate of cooling may vary but as a general guide a rate of between −1°C and −3°C per minute will prove suitable for the majority of cell cultures.
    23
  5. An alternative to the freezer system is the Taylor Wharton passive freezer where ampoules are held in liquid nitrogen vapour in the neck of Dewar. The system allows the ampoules to be gradually lowered thereby reducing the temperature. Rate controlled freezers are also available and are particularly useful if large numbers of ampoules are frozen on a regular basis.
  6. As a last resort if no other devices are available, ampoules may be placed inside a well-insulated box (such as a polystyrene box with sides that are at least 1 cm thick) and placed at −80°C overnight. It is important to ensure that the box remains upright throughout the freezing process. Once frozen, ampoules should be transferred to the vapour phase of a liquid nitrogen storage vessel and the locations recorded.
  7. If using a freezing method involving a −80°C freezer, it is important to have an allocated section for cell line freezing so that samples are not inadvertently removed. If this happens at a crucial part of the freezing process then viability and recovery rates will be adversely affected.
 
1.21 ISOLATION OF VIRUSES IN CELL CULTURE
The ability to culture viruses successfully in the laboratory depends on a number of important factors. These include the sensitivity of the cells used, the viability of the virus, the type of specimens sent to the laboratory and the way they are processed, the culture conditions and the stage of the illness when the specimen is taken. Even when all these considerations are taken into account, it must be remembered that not all viruses can be cultured and there are certain viruses that are very difficult to grow or require very specialized culture conditions. However, most of the more common pathogenic viruses can be cultured relatively easily provided proper conditions are satisfied. A wide variety of virus-sensitive cell lines are available either commercially or through one of the national or international cell bank collections such as the National Centre for Cell Sciences (NCCS), Pune, American the Type Culture Collection (ATCC, Rockville, Maryland, USA) or the European Collection of Animal Cell Cultures (ECACC, Salisbury, Wiltshire, UK). Cell culture systems used for isolation of different viruses are given in Table 1.2.
Cell cultures are normally grown in 25 cm2, 75 cm2 or 150 cm2 plastic tissue culture flascs depending on the volume of the cells required.
 
1.22 SUBCULTURING OF CELLS
 
Reagents and Equipment
  • Versene/trypsin solution
  • Growth medium
  • Fetal bovine serum, 10%
  • Sodium bicarbonate, 7.5%
    24
    Table 1.2   Cultivation of viral pathogens
    Virus family
    Easily isolated viruses
    Sensitive cell culture systems for isolation
    Preferred clinical specimens for viral isolations
    Herpes viridae
    Herpes simplex virus type 1 and 2
    African green monkey (VERO), Primary monkey kidney (PMK)
    Skin swab; genital swab
    Adenoviridae
    Adenovirus
    Human larynx carcinoma (Hep-2), Human embryonic kidney (HEK)
    Throat swab, feces, conjunctival swab
    Picornaviridae
    Echovirus
    Human diploid lung fibroblast (MRC-5), PMK, rhabdomyosarcoma (RD)
    Cerebrospinal fluid, stools
    Coxsackievirus type B
    PMK, VERO, RD
    Pericardial fluid, stools
    Polioviruses
    MRC-5, VERO
    CSF, stools
    Orthomyxoviridae
    Influenza A, B, C
    Madin-Darby canine kidney (MDCK), PMK
    Throat swabs; nasal swabs; nasopharyngeal aspirate
    Paramyxoviridae
    Parainfluenza 1, 2, 3, 4
    African green monkey kidney (LLC-MK2), PMK, MDCK
    Nasopharyngeal aspirate, nasal swab
    Measles
    Primary human embryonic kidney (PHEK)
    Reoviridae
    Rotavirus
    African green monkey kidney (BSC-1), Intestinal epithelium (CACO2)
    Stools
    Hepadenoviridae
    Hepatitis A
    Fetal rhesus kidney (FRK)
    Stools
    Retroviridae
    HIV
    T lymphocytes
    Blood
    Rhabdoviridae
    Rabies virus
    Hamster kidney cells; Chick embryo cells
    Urine, CSF, saliva; brain tissue
    Hepadenaviridae
    Hepatitis B virus
    Hepa RG cells
    Body fluids, blood
    Arboviruses
    Dengue, Japanese encephalitis
    C6/36
    Blood, throat swabs
    25
  • Cell culture flasks 75 cm2
  • Inverted microscope.
 
Procedure
  1. Examine the conditions of the cell monolayer using an inverted microscope and ensure that the cells are healthy and confluent.
  2. Discard the spent growth medium (GM) from the vessel and wash the monolayer twice with 10 ml of prewarmed versene/trypsin wash solution leaving the solution on the monolayer for 20 sec with each wash.
  3. Discard the wash solution and incubate the flask at 37°C for 2–5 min. Some cells are difficult to detach from the monolayer and require a more vigorous routine. In such cases, the cells may be incubated for longer intervals with the versene/trypsin solution left on.
  4. As the cells detach, add fresh prewarmed GM to the flask and pipette the cell suspension several times to break up the cell clumps.
  5. Count the cells and seed into fresh flasks, tubes or microtiter plates.
 
1.23 STANDARD VIRUS ISOLATION
  1. Seed 1 ml cell suspensions at a concentration of approximately 105 cells/ml into standard culture tubes using freshly made GM.
  2. Incubate the culture tubes in stationary racks at 37°C and allow to monolayer.
  3. When the monolayers are 90% confluent, discard the GM and replace with 4 ml MM per tube.
  4. Label the tubes accordingly and inoculate 0.2 ml freshly prepared specimen into each tube in duplicate.
  5. Keep at least two negative control tubes per rack.
  6. Incubate at 37°C. Some viruses (influenza and parainfluenza) need to be cultured at 33°C with trypsin but without FCS in the MM.
  7. Examine the cultures daily for CPE.
 
1.24 MICROTITER METHOD OF VIRUS ISOLATION
This method represents an enhancement of conventional monolayer isolation techniques. Using this method, six cell lines are seeded in suspension on microtiter plates thereby improving the sensitivity of virus isolation. Up to four specimens can be inoculated with each plate.
  1. Grow the selected cells in 75 cm2 plastic cell culture flasks in standard culture medium supplemented with 10% FBS and buffered with 7.5% sodium bicarbonate.
  2. Dispense confluent monolayers of cells by washing twice with equal volumes of a preheated versene/typsin mixture.
    26
  3. When the cells are confluent, replace GM with 100 μl MM.
  4. Make 10-fold serial dilutions of the virus isolate in clean sterile glass containers.
  5. Reconstitute the specific antiserum to a working concentration.
  6. Mix equal volumes (100 μl each) of virus and antiserum and incubate at 37°C for 1 hr. Make a positive control with virus and diluent only.
  7. Add 100 μl serum-virus mixture to each well and incubate at 37°C in a moist CO2 atmosphere.
  8. After 2–3 days examine the plate using an inverted microscope. A confirmed identification of the viral isolate is made when development of CPE has been effectively inhibited by the specific antiserum.
 
1.25 IMMUNOFLUORESCENCE (IF)
This procedure is very useful for confirming cell culture isolates and has been applied to a great effect in identifying a wide variety of viral antigens. It can also be used directly to detect viruses in clinical specimens. Two methods are used, direct and indirect. In the direct assay, infected cells are harvested, washed and fixed onto the wells of a Teflon-coated glass slide. The fixed cells are stained with a specific antiviral monoclonal antibody which is conjugated to a fluorescein dye Fluorescein isothiocyanate, (FITC). Unbound antibody is washed off and the slide is examined using a fluorescent microscope. The indirect method involves using an extra antibody and incubation step.
 
Reagents and Equipment
  • Versene/trypsin medium
  • PBS
  • Cold acetone
  • Mab-coupled FITC
  • Counting chamber
  • Incubator, 37°C
  • Fluorescent microscope.
 
Procedure
  1. Harvest the infected cells using a cell scraper or glass beads. Versene/trypsin may be used provided it does not interfere with the viral antigen.
  2. Spin the harvested cells gently at 300 to 400 g for 5 min.
  3. Discard the supernatant and resuspend the cells in PBS.
  4. Carefully count the cells using a disposable counting chamber and adjust the cell concentration to 1×106 cells/ml.
  5. Spot 15 μl of cell suspension onto the well of a Teflon-coated glass slide and air dry.
    27
  6. Fix the slide in cold acetone for 5 min at 4°C.
  7. After fixation, the slide is relatively stable and may be stored at 4°C for upto 24–48 hrs or frozen in a sealed container at −20°C to await staining at a later date.
  8. Add 20 μl of specific conjugate (monoclonal antibody tagged to FITC) to the well of the slide and incubate in a moist chamber at 37°C for 30 min.
  9. Wash the slide thoroughly for 10 min by immersing in a staining trough containing PBS.
  10. Allow the slide to air dry, mount in buffered glycerol and cover with a clean plastic cover slip.
  11. Read with ultraviolet light at 20–40X magnification.
 
1.26 QUANTITATION OF VIRUSES
Many approaches are available to determine the concentration of viruses in a given tissue. Infectivity assays, chemical assays and direct counting of virus particles using electron microscopy are among those used. The most widely utilized approach is the use of infectivity assays.
 
1.26.1 TCID50 (Tissue Culture Infective Dose)
This method determines the dilution of virus required to infect or cause CPE in 50% of inoculated cell cultures. The assay can be carried out in culture tubes or 96-well microtiter plates. A mouse fibroblast cell line showing characteristic CPE is shown in Figure 1.3.
  1. Make up a 50 ml cell suspension containing 1×105 cells/ml.
  2. Seed 1 ml of cell suspension into each of 50 sterile cell culture tubes and allow to monolayer by incubating at 37°C.
    zoom view
    FIGURE 1.3: L-929 Mouse fibroblast cell line showing characteristic CPE
    28
  3. When 90% of a cell monolayer has formed, the cultures are ready for viral inoculation. Prepare 10-fold (log) serial dilution of virus suspension in MM diluent (10−1 to 10−8).
  4. Remove the GM from each tube and replace with 1 ml of viral suspension. Inoculate five culture tubes for each virus dilution. Set up five control tubes which will contain MM diluent only.
  5. Incubate at 37°C.
  6. Monitor the cell cultures daily for signs of CPE.
The development of a CPE may be scored according to the following regime:
Grade of CPE
Percentage of cell monolayer infected
No CPE formed
±
< 30%
+
30 – 50%
++
50 – 75%
+++
> 75% of cell monolayer infected
Calculate the TCID50 by determining the dilution of virus causing CPE in 50% of the inoculated cell cultures (50% end point dilution). The appearance of any grade of CPE in the cell monolayer is indicative of infection.
Example of data used to calculate TCID50
Virus dilution
Infected cultures
Percent infected
10−1
5/5
100
10−2
5/5
100
10−3
4/5
91.7
10−4
4/5
91.7
10−5
2/5
37.5
10−6
1/5
10.0
10−7
0/5
0
 
Calculation of TCID50
The calculation is made according to the Reed-Muench method. It is clear from the above table that the 50% end point lies between the virus dilutions of 10−4 (77.7%) and 10−5 (34.5%). To find out exactly where the end point 29lies the proportionate distance between these two dilutions is first calculated.
zoom view
The 50% end point dilution is now calculated as follows:
zoom view
Since the virus inoculation was 1 ml per tube, the viral titer is therefore 104.7 TCID50/ml.
 
1.26.2 Plaque Assay
This is a focal assay which is based on the ability of infectious virus particles to form small areas of cell lysis or foci of infection on the cell monolayer. This is achieved by first adsorbing the virus onto a confluent cell monolayer and then overlaying the monolayer with agar. The overlay medium restricts the spread of secondary infection so that only areas of the cell monolayer adjacent to the initially infected cells will become infected and form plaques or small areas of CPE. These plaques can then be counted and the viral titer calculated. Plaque assays can be carried out in 24-well cell culture plates or Petri dishes (Fig. 1.4).
zoom view
FIGURE 1.4: Plaque assay
Serial dilutions of virus have been plated on confluent monolayer cultures of cells. The white areas show areas of the culture in which the cells have been killed. Each “plaque” is the result of the presence of one original infectious virus particle
30
 
Method
  1. Seed 2 ml of cell suspension containing 1×105 cells per ml into the wells of a 24-well cell culture plate and allow to form a healthy, confluent monolayer (2–3 days).
  2. Prepare overlay medium by combining 2x MM and agar.
  3. Equilibrate in a 50°C waterbath and ensure no clumping occurs.
  4. Prepare 10-fold (log) serial dilutions of virus suspension in MM diluent (10−1 to 10−8).
  5. Remove the GM from the wells of the culture plate.
  6. Inoculate duplicate wells with 0.2 ml virus suspension (two wells for each dilution) and absorb for 1 hr at 37°C. Tilt the plate at 15 min intervals to prevent the monolayer from drying out and to ensure an even distribution of inoculum.
  7. Wash the infected monolayers with prewarmed, sterile PBS to remove unabsorbed virus.
  8. Remove the overlay medium from the waterbath and allow to cool before pouring onto the monolayer. Ensure the overlay medium is not allowed to overcool or it will solidify prematurely.
  9. Carefully add 2 ml overlay into each well and incubate at 37°C to allow plaque formation.
  10. When plaques have formed, fix cell monolayers with 30% formaldehyde for 20 min.
  11. Carefully remove the overlay and stain the monolayer with 1% methylene blue solution.
  12. Remove the stain and count the number of plaques.
  13. Calculate the viral titer.
 
1.26.3 Calculation of Virus Titer from Plaque Assay
The infectivity of the virus is expressed as plaque-forming units (PFU) per ml and is calculated from the number of plaques observed at the appropriate dilutions.
Count the total number of plaques for each dilution, i.e. for each pair of wells since they have been inoculated in duplicate. From Table 1.3 it is clear that there are too many plaques to count at the lower dilutions.31
Table 1.3   Example of data used to calculate plaque titer
Dilution
Count 1
Count 2
Total count
10−1
> 100
> 100
> 200
10−2
> 100
> 100
> 200
10−3
49
72
121
10−4
30
36
66
10−5
15
9
24
10−6
3
5
8
10−7
0
0
0
10−8
0
0
0
The first countable plaques were therefore observed at a dilution of 10−3. Since the initial inoculum per well was 0.2 ml and the wells were inoculated in duplicate as follows:
0.4 ml
of a 10−3 dilution contains
121 PFU
0.04 ml
of a 10−3 dilution contains
66 PFU
0.004 ml
of a 10−3 dilution contains
24 PFU
0.0004 ml
of a 10−3 dilution contains
8 PFU
0.4444 ml
of a 10−3 dilution contains
219 PFU
Therefore, 0.4444 ml of undiluted virus contains 219 × 103 PFU = 2.19 × 105 PFU. Therefore, 1 ml of undiluted virus contains (1/0.4444) × 2.19 × 105 PFU = 4.93 × 105 PFU/ml.
 
1.27 TRYPSINIZING CELLS IN SERUM-FREE MEDIUM (SFM)
When trypsinizing cells in serum-free conditions, serum should not be used to inactivate trypsin as residues from the serum may be left in the cell suspension. All manipulations with cells should be carried out in a laminar flow cabinet. For best results, cells should still be in the log growth phase, i.e. not more than 80% confluent.
 
Reagents and Equipment
  • Trypsin/versene (2.5 mg/ml in PBS; Gibco)
  • Serum-free medium (SFM)
  • Tryspin inhibitor (TI) (Sigma)
  • Centrifuge
  • Inverted microscope
  • Hemocytometer
  • Sterile flasks
32
 
Procedure
  1. The day before trypsinization, cells should be fed with fresh medium.
  2. On the day of trypsinization, thaw trypsin, TI (in basal medium) and make up fresh stock of SFM. Do not warm trypsin solution.
  3. Remove medium from flask.
  4. Rinse cells with a small amount of trypsin and remove trypsin.
  5. Add in sufficient trypsin to cover the surface of the flask, a maximum of 0.5 ml per 25 cm2 flask.
  6. Allow flask to sit on bench for several minutes, monitoring the state of detachment regularly. As soon as the cells have rounded up (the length will depend on the cell type), gently tap the end of the flask to dislodge the cells.
  7. Add in an equal volume of TI.
  8. Remove cell suspension to a sterile universal.
  9. Wash flask with 5–10 ml sterile medium to remove residual cells and add to cell suspension.
  10. Centrifuge at 1,000 rpm for 5 min. If a pellet of cells has been formed, it may be necessary to centrifuge the cells for an additional 5 min.
  11. Very gently, remove the supernatant and resuspend the cells in 1 ml sterile medium. Add 9 ml medium.
  12. Centrifuge at 1,000 rpm for 5 min.
  13. Very gently, remove the supernatant and resuspend cells in 5 ml SFM.
  14. Determine the cell count and re-seed cells immediately.
 
1.28 PROTOCOL-9: TESTING FOR BACTERIAL AND FUNGAL CONTAMINATION
 
Aim
In cases of gross contamination the naked eye may identify the presence of bacteria and fungi. However, it is necessary to detect low-level infections by incubation of cell cultures and/or their products in microbiological broth. Equally these sterility tests can be used to confirm the absence of bacteria and fungi from the preparation which is important when preparing cell banks or cell culture products.
 
Reagents and Equipment
  • Soybean casein digest (Tryptone soy broth, TSB) (15 ml aliquots) TSB powder
  • Fluid thioglycolate medium (20 ml aliquots) (TGM)
    33
  • Bacillus subtilis
  • Candida albicans
  • Clostridium sporogenes
  • Personal protective equipment (latex medical gloves, laboratory coat, safety glasses)
  • Waterbath set to 37°C
  • Microbiological safety cabinet at appropriate containment level
  • Centrifuge
  • Incubator set at 32°C.
  • Incubator set at 22°C.
 
Procedure
  1. Culture cell line in the absence of antibiotics for 2 passages prior to testing.
  2. Bring attached cells into suspension with the use of a cell scraper. Suspension cell lines may be tested directly.
  3. Inoculate 2x Thioglycolate medium (TGM) and 2x Tryptone soy broth (TSB) with 1.5 ml test sample.
  4. Inoculate 2 (TGM) and 2 (TSB) with 0.1 ml C.albicans (containing 100 colony-forming units, CFU).
  5. Inoculate 2 (TGM) and 2 (TSB) with 0.1 ml B. subtilis (containing 100 CFU).
  6. Inoculate 1 TGM with 0.1 ml C. sporogenes (containing 100 CFU).
  7. Leave 2 (TGM) and 2 (TSB) un-inoculated as negative controls.
  8. Incubate broths as follows:
    • For TSB, incubate one broth of each pair at 32°C, the other at 22°C for 14 days
    • For TGM, incubate one broth of each pair at 32°C, the other at 22°C for 14 days
    • For the TGM inoculated with C.sporogenes. incubate at 32°C for 14 days.
  9. Examine test and control broths for turbidity after 14 days.
Criteria for a valid result: All positive control broths show evidence of bacteria and fungi within 14 days of incubation and the negative control broths show no evidence of bacteria and fungi.
Criteria for a positive result: Test broths containing bacteria or fungi show turbidity.
Criteria for a negative result: Test broths should be clear and show no evidence of turbidity.34
 
Note
  1. The positive controls should be handled in a laboratory remote from the main tissue culture laboratory.
  2. Control organisms (Bacillus subtilis, Clostridium sporogenes and Candida albicans) are also available from the national repositories.
  3. This test procedure should be carried out in a Microbiology laboratory away from the cell culture laboratory.
 
1.29 PROTOCOL-10: DETECTION OF MYCOPLASMA BY CULTURE
 
Aim
Detection of Mycoplasma by culture is the reference method of detection and has a theoretical level of detection of 1 colony-forming unit (CFU). However, there are some strains of Mycoplasma that are noncultivable (certain strains of Mycoplasma hyorhinis). The method is suitable for the detection of Mycoplasma in both cell cultures and cell culture reagents and results are obtained within 4 weeks. Mycoplasma colonies observed on agar plates have a ‘fried egg’ appearance.
 
Reagents and Equipment
  • 70% Ethanol in water
  • Mycoplasma pig agar plates (in 5 cm Petri dishes)
  • Mycoplasma pig agar broths (in 1.8 ml aliquots)
  • Mycoplasma synoviac
  • Mycoplasma pneumoniae
  • Personal protective equipment (sterile gloves, laboratory coat, safety visor)
  • Waterbath set to 37°C
  • Microbiological safety cabinet at appropriate containment level
  • CO2 incubator set at 32°C
  • Gas Jar (Gallenkamp)
  • Gas Pak Anaerobic System (Gallenkamp)
  • Gas Pak Catalyst (Gallenkamp)
  • Gas Pak Anaerobic Indicator (Gallenkamp).
 
Procedure
  1. Inoculate 2 agar plates with 0.1 ml of test sample.
  2. Inoculate 1 agar plate with 100 CFU M. pneumoniae.
  3. Inoculate 1 agar plate with 100 CFU M. synoviac
    35
  4. Leave 1 agar plate un-inoculated as a negative control.
  5. Inoculate 1 broth with 0.2 ml of test sample.
  6. Inoculate 1 broth with 100 CFU M. pneumoniae.
  7. Inoculate 1 broth with 100 CFU M. synoviac.
  8. Leave 1 broth tube un-inoculated as a negative control.
  9. Incubate agar plates anaerobically for 14 days at 37°C using a gas jar with anaerobic gas pak and catalyst.
  10. Incubate broths aerobically for 14 days at 37°C.
  11. Between 3 and 7 days and 10 and 14 days incubation, subculture 0.1 ml of test broth onto an agar plate and incubate plate anaerobically as above.
  12. Observe agar plates after 14 days incubation at 300X magnification using an inverted microscope for the presence of Mycoplasma colonies (see Figure 7.2, Chapter 7).
Criteria for a valid result: All positive control agar plates and broths show evidence of Mycoplasma by typical colony formation on agar plates and usually a colour change in broths. All negative control agar plates and broths show no evidence of Mycoplasma.
Criteria for a positive result: Test agar plates infected with Mycoplasma show typical colony formation.
Criteria for a negative result: The test agar plates show no evidence of Mycoplasma.
 
Notes
  1. Mycoplasma colonies have a typical colony formation commonly described as “fried egg” (see Fig. 3.3, Chapter 3) due to the opaque granular central zone of growth penetrating the agar surrounded by a flat translucent peripheral zone on the surface. However, in many cases only the central zone will be visible.
  2. Positive controls may be included at a concentration to give 100 colony-forming units. These controls should obviously be handled in a laboratory remote from the main tissue culture laboratory.
  3. Control organisms (M. pneumoniae, and M. synoviac) are available from National Collection of Type Cultures (UK), Institute of Microbial Technology Chandigarh or from any other national respository.
  4. Mycoplasma pneumoniae is a potential pathogen and must be handled in a class II microbiological safety cabinet.
  5. This test procedure should be carried out in a microbiology laboratory away from the cell culture laboratory.
36
 
1.30 PROTOCOL-11: TESTING FOR MYCOPLASMA BY INDIRECT DNA STAIN (HOECHST 33258 STAIN)
 
Aim
DNA staining methods such as Hoechst staining techniques are quick with results available within 24 hours, which compares favourably with 4 weeks for detection by culture. However, the staining of cultures directly with a DNA stain, results in a much-reduced sensitivity (~106 CFU/ml). This may be improved by co-culturing the test cell line in the presence of an indicator cell line such as Vero. This enrichment step results in a sensitivity of 104 CFU/ml of culture. This step also improves sensitivity by increasing the surface area upon which Mycoplasma can adhere. Like detection by culture, DNA staining methods are suitable for the detection of Mycoplasma from cell cultures or cell culture reagents.
 
Reagents and Equipment
  • Media—prewarmed to 37°C
  • 70% Ethanol in water
  • Methanol
  • Acetic acid glacial
  • Hoechst 33258 stain solution
  • Vero cells
  • Mountant (Autoclave 22.2 ml 0.2 M citric acid with 27.8 ml 0.2 M disodium phosphate. Add 50 ml glycerol. Filter sterilize and store at 4°C)
  • Mycoplasma hyorhinis
  • Personal protective equipment (sterile gloves, laboratory coat, safety visor)
  • Waterbath set to 37°C
  • Microbiological safety cabinet of appropriate containment level
  • Centrifuge
  • CO2 incubator set at 37°C
  • Microscope (UV Epi-fluorescence)
  • 35 mm plastic tissue culture dishes
  • Multidish 24-well
  • Cell scraper
  • Microscope slides and 22 mm cover slips
 
Procedure
  1. For each sample and control sterilize 2 cover slips in a hot oven at 180°C for 2 hours or by immersing in 70% ethanol and flaming in a blue Bunsen 37flame until the ethanol has evaporated. Also sterilize 2 coverslips to use as a negative control.
  2. Place the coverslips in 35 mm culture dishes (1 per dish).
  3. Store until needed.
  4. To prepare the Vero indicator cells add 2×104 cells in 2 ml of antibiotic-free growth medium to each tissue culture dish.
  5. Incubate at 37°C in 5% CO2 for 2 – 24 hrs to allow the cells to adhere to the coverslips.
  6. Bring attached test cell lines into suspension using a cell scraper. Suspension cell lines may be tested directly.
  7. Remove 1 ml of culture supernatant from duplicate dishes and add 1 ml of test sample to each. Inoculate 2 dishes with 100 CFU M. hyorhinis and 2 with 100 CFU M. orale.
  8. Leave duplicate tissue culture dishes un-inoculated as negative controls.
  9. Incubate dishes at 37°C in 5% CO2 for 1–3 days.
  10. After 1 day observe one dish from each pair for bacterial or fungal infection. If contaminated discard immediately. Leave the remaining dish of each pair for further 2 days.
  11. Fix cells to coverslip by adding a minimum of 2 ml of freshly prepared fixative (1:3 – glacial acetic acid: absolute methanol) to the tissue culture dish and leave for 3 to 5 minutes.
  12. Decant used fixative to toxic waste bottle. Add another 2 ml aliquot of fixative to cover slip and leave for a further 3 to 5 min. Decant used fixative to toxic waste.
  13. Air dry cover slip by resting it against the tissue culture dish for 30 to 120 min.
  14. Replace cover slip in dish and add a minimum of 2 ml Hoechst stain. Leave for 5 minutes shielded from direct light by aluminum foil.
  15. Decant used and unused stain to toxic waste.
  16. Add 1 drop of mountant to a pre-labeled microscope slide and place cover slip (cell side down) onto slide.
  17. Keep slide covered with aluminum foil, allowing it to set for at least 15 min at 37°C or for 30 min at room temperature.
  18. Observe slide under UV Epi-Fluorescence at 1000X.
Criteria for a valid result: Negative controls show no evidence of Mycoplasma infection. Positive controls show evidence of Mycoplasma infection Vero cells clearly seen as fluorescing nuclei.38
Criteria for a positive result: Samples infected with Mycoplasma are seen as fluorescing nuclei plus extra-nuclear fluorescence of Mycoplasma DNA (small cocci or filaments).
Criteria for a negative result: Uninfected samples are seen as fluorescing nuclei against a dark background. There should be no evidence of Mycoplasma.
 
Notes
  1. DNA stains such as Hoechst stain bind specifically to DNA. In all cultures cell nuclei will fluoresce. Uncontaminated cultures will show only fluorescent nuclei whereas Mycoplasma positive cultures contain small cocci or filaments which may or may not be adsorbed onto the cells.
  2. Hoechst stain is toxic and should be handled and discarded with care.
  3. Culture dishes should be placed in a sealed box or cultured in large Petri dishes to reduce evaporation.
  4. Positive cultures should obviously be handled in a laboratory remote from the main tissue culture laboratory.
  5. Control organisms (M. hyorhinis) are available from the National Collection of Type Cultures (UK). In India, they are available at the Institute of Microbial Technology, Chandigarh.